Ultrasound and glycosylation modifications enhance the physicochemical and functional properties of canola protein isolate for O/W emulsion stabilization

Abstract

This study aimed to prepare different modified canola protein isolates (CPIs), including ultrasound (U), glycosylation (G), ultrasound-glycosylation (U-G), and glycosylation-ultrasound (G-U). Assess the effect of different modifications on the quality characteristics of CPIs and the ability to stabilize emulsions. Sonication was found to increase the surface hydrophobicity resulting from the unfolding structure of CPI. The dual-modified U-G-CPI with 27.69 % glycosylation showed a solubility of 90.26 %, and an emulsification capacity of 80.87 m²/g. The emulsions stabilized by U-G-CPI exhibited higher degrees of protein adsorption (84.56 %) and a smaller particle size (4.22 μm) with a more homogeneous distribution at the oil-water interface. Additionally demonstrated enhanced thermal and salt stability resulting from its increased surface hydrophobicity and net charge. Overall, the ultrasound combined with glycosylation modification was found to significantly improve both the stability and emulsification properties of CPI, highlighting its potential as a plant-based emulsifier in the food sector and beyond.

Graphical abstract

Highlights

• •Ultrasound, glycosylation and dual modified CPI were prepared.
• •Ultrasound increased CPI structural flexibility and decreased CPI aggregation.
• •Dual modifications enhanced functional, especially emulsifying properties of CPI.
• •U-G-CPI stabilized emulsion has lower particle size and superior stability.

1. Introduction

In recent years, the demand for plant-derived protein sources has grown due to the increasing popularity of plant-based diets. Canola is widely cultivated throughout the world for oil production, second only to soya bean in terms of production. After extraction, the canola meal contains about 35–40 % protein, consisting mainly of salt-soluble 12S globulins (cruciferin, MW 300–350 kDa), and water-soluble 2S albumin (napin, MW 14–16 kDa) (Tang & Ghosh, 2021). Canola thus has significant untapped potential as a source of plant protein source. The nutritional value of canola is attributed primarily to its balanced composition of essential amino acids, including high levels of lysine, cysteine, and methionine, providing nutritional value comparable to that of animal proteins (Wanasundara et al., 2016). Proteins are especially useful as emulsifiers in foods as they can be adsorbed at the oil-water interface, resulting in the formation of an interfacial film. Unfortunately, protein emulsions tend to be unstable and susceptible to degradation by external factors, such as temperature, pH, and salt ion concentrations (Cui et al., 2023). Consequently, recent years have seen a significant focus on improving the emulsification properties of plant proteins, with the objective of achieving the desired performance.

To address the limitations of plant-derived proteins, various modification methods, including physical, chemical, and enzymatic approaches, are widely used to improve the emulsification properties of proteins (Tian et al., 2024). Physical modification involves the use of a force field to transform protein structures and alter their functionality. Ultra-high pressure and ultrasound are novel physical methods with the potential to enhance emulsification, but the costs associated with high-pressure modification have restricted their broader application (Goyal et al., 2024). Chemical modifications, including glycosylation, acylation, and other processes, involve a complex series of reactions. These reactions alter the multilevel structure, electrostatic charge, and hydrophobic groups of proteins, thereby influencing their interfacial properties (Glusac & Fishman, 2021). Although enzymatic treatment is an environmentally friendly and efficacious process, the prolonged reaction times required increase the risk of microbial contamination, restricting its industrial application and making it difficult to justify in terms of cost-benefit analysis (Fetzer et al., 2020).

Among protein modification methods, ultrasonication and glycosylation are considered to be the most promising and effective approaches. Ultrasound is a nonthermal processing technology that induces mechanical shear, cavitation, acoustic streaming, and localized heating effects (Ma et al., 2019). Previous studies have shown that these effects significantly decrease the particle size of the protein and enhance the functional properties of the protein isolate. However, detailed research on the changes in emulsifying properties has not yet been conducted (Jiménez et al., 2019). Glycosylation involves a chemical process in which sugar chains are covalently bonded to amino groups on protein molecules (Thirunavookarasu et al., 2022). Research conducted by Qu et al. (2018) showed that glycosylated rapeseed proteins had excellent emulsifying properties, accompanied by improved solubility, gelation, and thermal stability. Similar results were found in soy proteins (He et al., 2023). Proteins can form complexes with sugars, increasing molecular mass and exposing the internal hydrophobic groups, facilitating adsorption at the oil-water interface and thus improving emulsion stability.

Recent research has explored various modifications to enhance protein function. However, certain limitations were observed using single-modification techniques. On this basis, therefore, the use of dual modification has been proposed to achieve synergistic effects (Zhang, Wang, et al., 2023). These techniques have included the combining of glycosylation with succinylation (He et al., 2023), and glycation with heating (Wang et al., 2019), improving both protein emulsification activity and stability. Yang et al. (2023) demonstrated that ultrasound-assisted flaxseed gum (FG) glycosylation enhanced the interfacial properties of pea protein (PP). The double modified PP emulsions showed smaller droplet sizes, superior thermal stability, with good stability observed during the storage over 42 days. These improvements are attributed to FG's unique molecular structure as an anionic polysaccharide, which forms complexes with proteins (Liu et al., 2018), thereby modulating surface hydrophobicity and improving emulsification efficiency. Additionally, FG offers health benefits, including reducing the incidence rate of diabetes and preventing colorectal cancer (Yang et al., 2023). Despite these advantages, few studies were carried out to investigate the effect of FG on improving the functional properties of CPI.

Under the above situation, our study endeavors to comprehensively evaluate the effects of ultrasonication and flaxseed gum glycosylation (both single and dual modifications in different orders) on the structural, physicochemical, and functional properties of CPI. Furthermore, different modified proteins were applied to stabilize emulsions, and the emulsion properties were investigated from their particle size, ζ-potential values, microstructure, and storage stability, thereby providing new ideas to broaden the application of canola protein as an emulsifier in the food industry.

2. Materials and methods

2.1. Materials

Canola protein isolate (CPI, purity >95.0 %) and flaxseed gum (FG, purity >98.0 %) were purchased from Henan Boyan Biological Technology Co., Ltd. (Zheng Zhou, China). Flaxseed oil was extracted from the samples using an Xz-z505w horizontal oil screw press (Guangzhou Xuzhong Food Machinery Co., Ltd., China). All chemical reagents were provided by Macklin Biochemical Co., Ltd. (Shanghai, China). Deionized (DI) water was used in all experiments.

2.2. Protein modifications

CPI powder (1.0 wt%) and FG powder (0.2 wt%) were dissolved in DI water with continuous dispersion for 2 h. Samples with different modifications of CPI were recorded as U-CPI, G-CPI, U-G-CPI, and G-U-CPI, representing modifications induced by ultrasound, glycosylation, ultrasound followed by glycosylation, and glycosylation followed by ultrasound, respectively.

2.2.1. Ultrasonication

U-CPI was obtained by ultrasonic treatment of CPI by using a 20 kHz ultrasonic homogenizer (JY92-IIDN, NingboScientz Biotechnology Co., Ltd., China). The prepared CPI solutions were stored in 500 mL beakers. Then, a 20 mm diameter ultrasonic probe was inserted into the solution, positioned 2 cm above the bottom. The CPI dispersion was sonicated at an ultrasonic power of 480 W, ultrasonic density of 0.96 W/mL for 20 min (5 s:2 s, work/rest cycles). To prevent protein denaturation, the sonication process was carried out in an ice-water bath.

2.2.2. Glycosylation

Based on preliminary experiments, G-CPI was obtained by mixing the CPI and FG solutions with a ratio of 6:1 (v/v) at 70 °C with continuous stirring for 4 h. Thereafter, the reaction was halted by cooling in an ice-water bath. To remove unreacted FG and proteins, the reaction mixture was dialyzed against ultrapure water at 4 °C for 72 h using a dialysis membrane with a molecular weight cut-off (MWCO) of 8–14 kDa and a flat width of 77 mm.

2.2.3. Dual modifications

The reaction conditions were optimized through preliminary tests comparing grafting efficiency. U-G-CPI was prepared by ultrasonication of CPI at ultrasonic power of 480 W, ultrasonic density of 0.96 W/mL for 20 min (5 s:2 s, work/rest cycles), followed by the addition of FG at a ratio of 6:1 (v/v) and continuous stirring at 70 °C for 4 h. By contrast, G-U-CPI was obtained by mixing CPI and FG solutions, followed by immediate sonication.

2.3. Degree of glycosylation

The degree of grafting (DG) was calculated via the o-phthaldialdehyde (OPA) assay. The OPA solution was freshly prepared according to the method described by He et al. (2023). Two hundred microliters of 0.4 % sample solutions were mixed with 4  mL of OPA reagent and incubated at 35 °C for 2 min. The absorbances were measured at 340  nm using a UV spectrophotometer (TU-1780, Shimadzu, Kyoto, Japan). The DG values of the G-CPI, U-G-CPI, and G-U-CPI conjugates were calculated as follows:

$$\mathit{DG}\left(\%\right)=\frac{A_{\mathit{0}}-A_{\mathit{1}}}{A_{\mathit{0}}}\times 100\%$$

where A₀ and A₁ are the absorbance values of CPI before and after glycosylation in different orders, respectively.

2.4. Physicochemical and structural characterization of modified CPI

2.4.1. Free sulfhydryl content

Tris-glycine buffer (pH = 8.0) was prepared with 0.086 mol/L Tris, 0.09 mol/L glycine, and 0.4 mmol/L EDTA. Ellman's solution was prepared by dissolving 120 mg of Ellman's reagent (5,5′-dithiobis(2-nitrobenzoic acid, DTNB) in 30 μL of the buffer. Samples (1 mg/mL) were dissolved in 50 μL of Ellman's solution, incubated for 40 min at room temperature, and centrifuged (10,000 g, 8 min). The absorbance of supernatant was measured at 412 nm. The content of free sulfhydryl group was calculated according to the following formula:

$$\mathit{Free}\mathit{SH}\left(\mathit{\mu mol}/g\right)=\frac{73.53\times A\times D}{C}$$

where A is the absorbance of the samples at 412 nm, C is the sample concentration (mg/mL) and D is the dilution ratio.

2.4.2. Surface hydrophobicity (H₀)

As described by Liu et al. (2023), CPI sample solutions with different concentrations (0.02–0.1 mg/mL) were prepared in 0.01 mol/L phosphate buffer (pH = 7.4). Each sample (5 mL) was mixed with 20 μL of 8 mmol/L 8-anilinonaphthalene-1-sulfonate (ANS) solution for 120 s, and fluorescence intensity was measured using a fluorescence spectrometer (RF-530, Shimadzu, Japan) at excitation and absorption wavelengths of 390 nm and 470 nm, respectively. The linear slope of the plot of fluorescence intensity versus protein concentration represented the surface hydrophobicity.

2.4.3. Fourier-transform infrared (FTIR) spectroscopy

The FTIR spectra of protein samples were determined using a Fourier Transform Infrared Spectroscopy (FTIR, Nicolet 6700, Shimadzu, Japan). All the samples were blended with KBr in a 1:100 ratio, scanned and recorded from 400 to 4000 cm⁻¹, with a resolution of 4 cm⁻¹ and an accuracy of 0.01 cm⁻¹. The software “Peakfit Version 4.12” was used to analyze the secondary structure of the protein (Cui et al., 2020).

2.4.4. Intrinsic fluorescence spectroscopy

Protein samples (150 mg) were dissolved in phosphate buffer (0.01 mol/L, pH = 7.0). Impurities were removed by filtration using a water separation membrane. Emission spectra over the 300–500 nm range were scanned with a fluorescence spectrophotometer, using an excitation wavelength of 295 nm, a slit width of 5.0 nm, and a scanning speed of 5 nm/s (Cui et al., 2020).

2.4.5. Ultraviolet-visible (UV–vis) spectroscopy

Protein samples (150 mg) were dissolved in phosphate buffer (0.01 mol/L, pH = 7.0). Analyzed by a UV–vis spectrophotometer (UV-2400, Shimadzu, Kyoto, Japan) at a wavelength of 200–800 nm.

2.4.6. Scanning electron microscopy

The surface morphology of protein samples was evaluated using scanning electron microscopy (SEM, JSM-6610LV, Hitachi, Japan). Samples were applied with a cotton swab, fixed on glass slides, coated with gold, and scanned at 1000 times magnification.

2.5. Functional properties of modified CPI

2.5.1. Solubility

Sample solubility was determined using the centrifuge method as previously described (Liu et al., 2023). Protein solutions (10 mg/mL) were prepared in phosphate buffer and centrifuged at 8000 r/min for 10 min. Protein concentrations were measured by the Lowry method and expressed as a percentage of solubility.

2.5.2. Water and oil holding capacity (WHC/OHC)

WHO and OHC were evaluated by the method described by Patil et al. (2024). Protein samples were weighed 1 g in the pre-weighed centrifuge tubes. For WHC, 10 mL of DI water was added, while 10 mL of soybean oil was added for OHC. Maintained for 30 min, the tubes were centrifuged for 10 min at 8000 r/min, and the released water and oil were drained. WHC and OHC were calculated with the following equations.

$$\mathit{WHC}/\mathit{OHC}\left(g/g\right)=\frac{m_2-m_1}{m_0}$$

where m₀ is the initial mass of the sample, m₁ and m₂ represent the mass of protein sample pre- and post- centrifugation, respectively.

2.5.3. Foaming properties

Foaming capacity (FC) and foam stability (FS) were calculated as described by Cui et al. (2023). A volume of 15 mL (1 mol/L) of the sample was placed in a measuring cylinder (V₁), followed by homogenization at 12,000 rpm for 2 min using a high-speed disperser (FJ300-SH, Shanghai, China), and the foam volume (V₂) was recorded. After standing for 30 min, the foam volume (V₃) was recorded. FC and FS were calculated with the following equations.

$$\mathit{FC}\left(\%\right)=\frac{V_2}{V_1}\times 100\%$$

$$\mathit{FS}\left(\%\right)=\frac{V_3}{V_2}\times 100\%$$

2.5.4. Emulsibility and emulsifying stability

Emulsifying activity index (EAI) and emulsifying stability index (ESI) were measured by referring to the method of Lisuzzo et al. (2022). Twenty milliliters of sample solutions (1 mg/mL) were mixed with soybean oil in a 3:1 ratio and homogenized at 7000 r/min for 2 min, after which 100 μL of the emulsion was pipetted from the bottom mixed with 5 mL of SDS solution (pH = 7.0, 0.1 %), and the absorbance value was measured by UV spectrophotometer at 500 nm. The same treatment was repeated after 20 min. EAI and ESI were calculated with the following equations.

$$\mathit{EAI}\left(m^2/g\right)=\frac{2\times 2.303\times A_0\times N}{\phi \times L\times C\times 10000}$$

$$\mathit{ESI}\left(\min \right)=\frac{A_0\times 20}{A_0-A_{20}}$$

where N represents the dilution coefficient (100), φ is the fraction of the oil phase (φ = 0.25), L is the optical path (L = 1 cm), C represents the concentration of protein solution (g/mL), A₀ and A₂₀ denote absorbance of the sample at 0 and 20 min, respectively.

2.6. Preparation of flaxseed oil emulsions

The flaxseed oil emulsions were prepared by our preliminary experiments. Specifically, the CPI, U-CPI, G-CPI, U-G-CPI, and G-U-CPI (1 wt%) dispersions described in Section 2.2 were each mixed with flaxseed oil (3:1, v/v). The mixtures were then dispersed using a high-speed disperser at a speed of 8000 rpm for 10 min. Subsequently, the initial emulsion was subjected to a high-pressure homogenizer (SAMRO Co., Ltd., China) with four treatments at 20 MPa to obtain stabilized emulsions. Proclin 950 (0.02 %, m/m) was incorporated into all emulsions as an antimicrobial preservative.

2.7. Droplet size and ζ-potential of emulsions

As described by Diao et al. (2024), the particle size of the dispersed emulsion was determined using a laser diffraction instrument (Mastersizer 2000, Malvern Instruments Co., Ltd., UK). The refractive indices of the dispersing medium and protein particles were 1.33 and 1.46, respectively. Meanwhile, a Zeta sizer Nano recorded the ξ-potential of different emulsions (ZS90, Malvern Instruments, Ltd., UK).

2.8. Adsorbed protein of emulsions

The percentages of interface-adsorbed proteins (AP%) of emulsions were determined based on the method of Diao et al. (2024). Two milliliters of emulsion were centrifuged at 10,000 rpm for 60 min, and the supernatant was filtered through a 0.45 μm filter. The content of adsorbed proteins were quantified using the Lowry method.

2.9. Apparent viscosity of emulsions

The apparent viscosity of the emulsions was measured using a rotational rheometer (HAAKE MARS60, Thermo Scientific, USA) with 40 mm parallel-plate geometry. The gap between plates was set to 1 mm, and measurements were taken at 25 °C with shear rates ranging from 0 to 100 s⁻¹.

2.10. Microstructural characterization of emulsions

Confocal scanning laser microscopy (CSLM) was obtained using a Nikon confocal microscope (A1R+, Nikon, Japan). A solution of 0.1 % Nile Red in isopropanol was used to stain the oil phase of 0.5 mL of emulsion, and observations were made at an excitation wavelength of 458 nm.

2.11. Emulsion stability against environmental stresses

2.11.1. Thermal stability

Freshly prepared emulsions were heated at 90 °C for 30 min, and changes in particle size were observed before and after treatment to assess the thermal stability.

2.11.2. Saline ion stability

To evaluate the salt ion stability of the emulsions, the samples were mixed with equal volumes of NaCl solutions of different concentrations (100 and 300 mM), followed by overnight incubation at 25 °C and measurement of the particle size.

2.12. Emulsion storage stability

Freshly prepared emulsions were placed in a refrigerator at 4 °C for 14 days without light. Their particle sizes were recorded on days 0, 7, and 14. Before measuring particle size, all emulsions were gently mixed to ensure homogeneity. Samples were then taken from the middle layer using a pipette and immediately analyzed.

2.13. Statistical analysis

Each experiment was performed in triplicate, and the results are expressed as mean ± standard deviation. One-way ANOVA and Tukey's test were used for statistical comparisons. Values of P < 0.05 were considered statistically significant. All the analysis was conducted using IBM SPSS 26.0 (IBM Corp., Armonk, NY, USA).

3. Results and discussion

3.1. Degree of glycosylation

Glycosylation represents the primary stage of the Maillard reaction, and the degree of glycosylation is associated with the loss of active amino groups (Sun et al., 2020). Thus, the extent of the Maillard reaction can be quantified by measuring the amount of free amino groups and characterized by the DG. As shown in Table 1, the DG value of G-CPI was 15.76 ± 1.24 %. With the superimposition of ultrasonic treatment, the DG values of U-G-CPI and G-U-CPI were significantly increased to 27.69 ± 1.05 % and 20.93 ± 1.13 %, respectively. Ultrasound-induced cavitation effects induce conformational changes in CPI, enlarging the areas of contact between CPI and FG, and promoting the formation of additional covalent bonds, thus enhancing the reaction with carbonyl groups on the surfaces of polysaccharide molecules (Li, Wang, et al., 2022). Moreover, ultrasonication before glycosylation alters the spatial conformation of the protein, exposing more ε-amino acid lysine residues, leading to an increased interaction between the protein and polysaccharide molecules (Li, Xi, et al., 2022).

Table 1.

The degree of glycosylation of G-CPI, U-G-CPI, and G-U-CPI.

Sample	G-CPI	U-G-CPI	G-U-CPI
DG (%)	15.76 ± 1.24b	27.69 ± 1.05a	20.93 ± 1.13a

Note: Glycation-canola protein isolate (G-CPI), Ultrasound-glycation-canola protein isolate (U-G-CPI), Glycation-ultrasound-canola protein isolate (G-U-CPI), Different lowercase letters indicated significant differences (P < 0.05).

3.2. Physicochemical properties of modified CPI

3.2.1. Surface hydrophobicity (H₀)

Surface hydrophobicity can be considered an indicator of the number of hydrophobic groups present on the protein surface, which has a substantial impact on properties such as solubility and emulsification (Li, Zhang, et al., 2022). As shown in Fig. 1A, ultrasound significantly increased the H₀ value of CPI from 723.3 to 951.65 (P < 0.05), attributed to the induction of protein unfolding and macromolecular breakdown by the cavitation effect. Similar findings were reported by Tian et al. (2023) for phycocyanin, where ultrasound treatment led to exposure of hydrophobic groups, thus increasing the surface hydrophobicity. Following glycosylation, H₀ decreased to 577.2 due to the incorporation of additional groups (-OH) during the glycosylation process, which increased the density of hydrophilic moieties on the protein surface. Dual modification further reduced to 497.3 (U-G-CPI) and 526.6 (G-U-CPI). Sonication facilitated the glycosylation reaction by exposing internal groups of the protein, accompanied by partial masking of some surface hydrophobic regions by the attached sugar molecules during the glycosylation process. However, this balances the protein function by increasing the solubility and stabilizing the emulsion (Spotti et al., 2019). Additionally, the introduced hydroxyl groups by glycosylation resulted in a decline in the amount of free lysine, thus reducing the number of ANS binding sites (Sun et al., 2020). Post-glycosylation sonication may result in an extension of the polypeptide chain. However, steric hindrance resulting from FG-induced exposure to hydrophobic groups also limited ANS accessibility and led to a marked decrease in surface hydrophobicity.

Fig. 1.

H₀ (A) and free-SH content (B) of CPI with different modifications.

3.2.2. Free sulfhydryl (–SH)

Fig. 1B shows the content of free sulfhydryl groups in CPI following different treatments. There is no doubt that ultrasound treatment led to the exposure of a considerable number of internal free sulfhydryl groups on the surface of the protein molecule, and significantly increased the free sulfhydryl of U-CPI content from 5.00 to 14.31 μmol/g. Due to the cavitation and mechanical effects of ultrasound leading to disruption of the protein structure, the surface exposure of internal sulfhydryl groups, together with the breakage of disulfide bonds, led to increased free sulfhydryl contents. The presence of free sulfhydryls is crucial for the creation of new disulfide bonds during protein–protein interactions. However, our findings indicated a reduction in free sulfhydryl groups in CPIs following dual modification. In U-G-CPI, the content slightly decreased, attributed to the effects of glycosylation accelerated after ultrasound, where sulfhydryl groups reacted with protein molecules or polysaccharides to form thioether bonds and other conjugates (Li, Zhang, et al., 2022). In G-U-CPI, the single glycosylation step stabilized the protein structure, limiting sulfhydryl exposure, while subsequent sonication disrupted disulfide bonds and thus increased the sulfhydryl content. Furthermore, partial oxidation of sulfhydryl groups by hydrogen peroxide generated by sonication cavitation would also to the process (Sun et al., 2020; Tian et al., 2023). Similar results were observed in a study by Chen, Dai, et al. (2022) on pea protein isolate (PPI), where ultrasound and glycosylation increased both H₀ and -SH content. However, increased temperature promoting free radical generation would induce oxidation of sulfhydryl groups resulting in the formation of disulfide bonds.

3.2.3. Fourier-transform infrared spectroscopy (FT-IR)

FTIR is a crucial analytical technique that can reflect alterations in chemical bonds and groups within protein molecular structures. The FTIR spectra of CPI with different modifications (Fig. 2A) showed that sonication did not lead to the introduction of new peaks or the attenuation of intrinsic peaks. However, G-CPI, U-G-CPI, and G-U-CPI showed minor alterations in the amide I and II regions, indicating changes in the secondary structure of the protein. Precisely, the emergence of a novel peak at 1550 cm⁻¹ indicated that glycosylation affected the surrounding amino acid milieu, thereby altering N—H bending and C—N stretching vibrations (Badar et al., 2024). The emergence of this peak may reflect the introduction of new vibrational modes from hydroxyl groups (—OH) and other groups, which form a hydrogen bonding network, and modify the protein conformation and thus intermolecular hydrogen bonding. It was also observed that the dual sonication and glycosylation modification resulted in stronger absorption in the 3700–3200 cm⁻¹ range, suggesting stretching of the spatial structure. The increased exposure of non-polar groups (such as the hydrophobic groups) in U-G-CPI, driven by the cavitation effect promoted glycosylation and the formation of covalent N—O bonds, further altering the protein secondary structure (Boostani et al., 2017).

Fig. 2.

FT-IR spectra (A), second structure composition (B), intrinsic fluorescence emission spectra (C), and UV–Vis spectra (D) of CPI with different modifications.

Changes in CPI's secondary structure were analyzed using Peakfit software (Fig. 2B). The natural CPI comprised 20.61 % α-helix, 25.96 % β-sheet, 36.84 % β-turn, and 16.59 % random coils. After ultrasonic treatment, α-helix content decreased to 12.59 %, while the proportion of random coils increased, indicating further disruption of the ordered protein structure by the ultrasonic treatment. Glycosylation increased the β-turn content to 33.06 %, suggesting alterations in the CPI secondary structure with partial unfolding of the molecule. These results are consistent with those of Qu et al. (2018) showing that protein secondary structure may be affected due to protein–polysaccharide interactions as well as intermediates of the Maillard reaction (He et al., 2023). Notably, the consistent reduction in α-helix content in U-G-CPI and G-U-CPI suggests that these samples would exhibit enhanced protein–protein interactions, thus contributing to the stability of the protein film at the oil-water interface (Zhang, Zhao, et al., 2023). Potentially, the binding of dual-modified CPI to the oil surface is favourable, suggesting an improvement in emulsification properties.

3.2.4. Fluorescence spectroscopy

Fluorescence emission reflects changes in protein conformation due to alterations in the polarity surrounding tryptophan (Trp) residues. Natural CPI showed fluorescence absorption at λ_max = 340 nm, indicating that Trp residues were situated in a polar environment (Zhang et al., 2014). The increased fluorescence absorption intensity at λ_max = 340 nm observed following ultrasonic treatment indicates that cavitation-induced shear and molecular collisions exposed buried chromophores, leading to the enhanced fluorescence intensity of CPI. Glycosylation resulted in a reduction in the fluorescence intensity at the same wavelength, which was primarily attributed to the binding of hydrophilic groups to fluorescent chromophores, thereby gradually exposing Trp and tyrosine residues to polar environments and lowering the endogenous fluorescence (Chen et al., 2019; Tian et al., 2024). Interestingly, our study revealed that altering the order of the modifications resulted in an initial increase in the increased fluorescence intensity. However, at a certain point, the internal chromophore was no longer exposed due to aggregation between protein molecules, leading to a reduction in fluorescence intensity compared to the natural CPI. Additionally, the λ_max of U-G-CPI and G-U-CPI exhibited a 5 cm blue shift, indicating a shift of the amino acid environment toward a more hydrophobic state (Qu et al., 2018). Under the synergistic effects of ultrasound and glycosylation, hydrophobic interactions and protein–protein hydrogen bonding dominated rather than protein–polysaccharide interactions.

3.2.5. UV–vis spectroscopy

To further characterize the conformational changes in the tertiary structure of CPI, UV spectra were obtained in the range of 180–340 nm. It is widely accepted that protein absorption near 225 nm is due to peptide bonds (—CO—NH—), while absorption near 270–280 nm corresponds to Trp and tyrosine (Tyr) residues. Fig. 2D clearly distinguished the UV spectra of modified CPIs. The increasing UV absorbance in U-CPI following sonication can be attributed to the unfolding of the molecular structure of the protein aggregates under strong shear forces, exposing greater numbers of Trp and Tyr residues and enhancing UV absorption. Interestingly, ultrasonic modification extended the structure of CPI, with the addition of flaxseed gum facilitating a harmonious Maillard reaction, involving amino acids such as tyrosine, histidine, and phenylalanine, further enhancing UV absorption (Li, Xi, et al., 2022).

In addition, the trend seen in the fluorescence spectroscopy results was observed, in which glycosylation followed by sonication resulted in a reduction in UV intensity due to the excessive shear forces, promoting electrostatic interactions between the protein surfaces leading to aggregation (Zhang et al., 2021). The results suggested that the tertiary structure of CPI can be loosened following the dual modification, which may facilitate its dispersion in an aqueous solution.

3.2.6. Scanning electron microscopy (SEM)

The microstructure of CPI and the samples treated with different methods are visualized by SEM analysis. The morphology of the CPIs, as depicted in Fig. 3, unmodified CPI appears as discrete, roughly spherical aggregates. In contrast, the modified samples exhibit irregular, sheet-like structures, partly as a consequence of freeze-drying and the effects of sonication and glycosylation. The dual modifications resulted in the appearance of a more open and disordered protein structure, consistent with the FT-IR results presented in Section 3.2.3. Additionally, previous studies have demonstrated the disintegration of proteins into small colloidal particles through sonication and chemical treatments, such as glycation-heat (Wang et al., 2019) and glycosylation (Tian et al., 2024). In conclusion, the dual treatment of sonication and glycosylation significantly altered the protein structure, correlating with changes in the functional properties.

Fig. 3.

SEM images of the morphological structures of different modified CPI. CPI (A), U-CPI (B), G-CPI (C), U-G-CPI (D), and G-U-CPI (E).

3.3. Functional properties of modified CPI

3.3.1. Solubility

Solubility represents the degree to which protein molecules disperse in solution, remaining unprecipitated or aggregated (Boostani et al., 2017). It is a key indicator affecting other functional properties; thus, we evaluated the solubility of CPI with different modifications (Fig. 4A). The solubility of U-CPI increased from 57.53 % for native CPI to 65.19 % at pH = 7, as ultrasonic treatment induced partial unfolding of protein molecules and enhanced protein–water interactions (Jiménez et al., 2019). Following glycosylation and dual modifications, the solubility further improved to 72.86 % (G-CPI), 90.26 % (U-G-CPI), and 83.65 % (G-U-CPI). The introduction of ultrasound notably increased the degree of glycosylation of CPI, resulting in a higher number of hydrophilic -OH groups in the conjugates compared to the unmodified protein (Qu et al., 2018). Moreover, the reduced surface hydrophobicity observed in Section 3.2.1 further confirms that an increased grafting degree leads to enhanced solubility. In addition, as reported by Jiménez et al. (2019) and Patil et al. (2024), the synergistic effect of ultrasound and glycosylation not only promotes glycosylation but also reduces protein aggregation through partial unfolding and exposure of reactive sites, thereby improving solubility. Our findings are consistent with the study of Chen, Zhao, et al. (2022), who observed a 70 % increase in solubility after sonication of PPI-arabinose conjugates.

Fig. 4.

Solubility (A), WHC and OHC (B), FC and FS (C), EAI and ESI (D) of CPI with different modifications.

3.3.2. Water- and oil-holding capacity (WHC/OHC)

Interactions between water, oil, and proteins in food systems are of great importance and significantly impact on the flavors and organoleptic properties (Jiménez et al., 2019). WHC refers to the ability of a protein to retain water, as shown in Fig. 4B, the WHC increased significantly in all CPI samples after sonication and glycosylation. Notably, the WHC of G-U-CPI exhibited a significant rise (P < 0.05) from 2.97 ± 0.03 g/g to 4.96 ± 0.05 g/g. The introduction of hydrophilic sugar molecules facilitates hydrogen bonding with water molecules, while subsequent sonication exposed more hydrophilic groups thus increasing the surface area for interaction with water, and producing a synergistic effect that enhances the WHC.

OHC, indicating the oil-binding ability of protein aggregates, also increased after modification (Fig. 4B). The OHC of U-CPI and G-CPI increased by 0.38 and 0.41 g/g, respectively. This is a consequence of an alteration in the protein structure that enhanced physical interaction with oil. After dual modification, the OHC significantly increased to 4.22–4.54 g/g, consistent with the Ho trend shown in Fig. 1B and similar to the observations reported on ovalbumin modified by succinylation and sonication (Zhang, Zhao, et al., 2023). In contrast to the WHC results, U-G-CPI showed superior OHC, likely due to the reduced impact of sonication on OHC, thereby enhancing the stability of the protein matrix and improving oil adsorption.

3.3.3. Foaming capacity and stability (FC and FS)

Foaming capacity of protein is the extent to which the protein adsorbs bubbles at the oil/water interface (Chen, Dai, et al., 2022). As shown in Fig. 4C, both FC and FS were significantly increased in U-CPI and G-CPI compared to CPI (P < 0.05). The enhanced FC of U-CPI can be attributed to the mechanical force and cavitation effect generated by ultrasonication, breaking the protein aggregates into smaller, dispersed structures and increasing the overall surface area (Li, Zhang, et al., 2022), thereby enhancing the FC. This provided a superior enhancement of FC in comparison to that of the glycosylation modification. In contrast, G-CPI showed better FS due to peptide chain elongation during glycosylation, which strengthens hydrophobic interactions at the air-water interface and promotes stronger interactions between hydrophobic residues than seen in the natural state (Chen, Zhao, et al., 2022). Dual modifications in different orders resulted in a notable increase in FC, from 32.71 ± 0.39 % to 80.00 ± 1.79 % (U-G-CPI) and 85.42 ± 1.00 % (G-U-CPI), with increases in FS from 62.5 ± 1.25 % to 86.88 ± 0.97 % (U-G-CPI) and 93.4 ± 1.97 % (G-U-CPI). The combined effect of ultrasound and glycosylation loosened the protein structure, facilitating rapid adsorption at the air-water interface and allowing the formation of a stable foam layer (Cui et al., 2023). Notably, G-U-CPI exhibited superior FC and FS, where ultrasonication enhanced protein unfolding, facilitating glycosylation and preventing foam collapse by creating a firmer protein matrix, thus enhancing its overall stability (Kostova, 2023).

3.3.4. Emulsibility and emulsifying stability

Emulsibility represents the ability of proteins to form stable emulsions by interacting with water and oil, and plays a crucial role in emulsion stability. As depicted in Fig. 4D, both single and dual modifications significantly improved the EAI of CPI (P < 0.05). After the dual modifications, EAI increased from 45.65 m²/g in unmodified CPI to 80.87 m²/g for U-G-CPI and 76.69 m²/g for G-U-CPI. Ultrasonication promotes protein denaturation, and this structural disruption enhances protein adsorption at the oil-water interface (Jiménez et al., 2019). It has been widely acknowledged that the most significantly enhanced attribute associated with protein–polysaccharide complexation is the emulsification capacity, while glycosylation enhanced hydrophilicity through the incorporation of –OH groups (Qu et al., 2018; Zhang, Wang, et al., 2023) Furthermore, both the FT-IR and SEM findings have corroborated that the presence of an extended secondary structure and more permeable surface facilitated rapid adsorption, which in turn enhances the EAI. Notably, the U-G-CPI has a higher emulsifying activity, due to the action of ultrasound in promoting the rapid adsorption of proteins at the oil-water interface and reducing the interfacial tension. This results in an equilibrium between hydrophilic and hydrophobic moieties on the surface of the molecule (Chen, Dai, et al., 2022), this outcome aligns with the OHC results.

As expected, different modifications significantly improved the ESI of CPI (P < 0.05), with the ESI for both U-G-CPI and G-U-CPI increased by a factor of 1.39, as glycosylation enhanced the interfacial properties of the emulsifier to prevent droplet aggregation (Lisuzzo et al., 2022), while the turbulence behaviour induced by ultrasound leads to a significant reduction in the droplet size, promoting a more homogeneous distribution of the dispersed phase (Patil et al., 2024). The combination of glycosylation and ultrasonic modifications thus plays an important role in CPI emulsification activity and emulsion stability, similar results were reported by Cui et al. (2020) with improved soybean protein isolate emulsification after succinylation and sonication.

To further investigate the effects of different modifications on the emulsifying properties of CPI and its stabilization potential, further investigation of the emulsion performance of both single and dual-modified CPI based on the comprehensive results mentioned above was undertaken.

3.4. Droplet analysis and interfacial characteristics of emulsions

3.4.1. Particle size and ζ-potential

To provide information on the effects of sonication and glycosylation on particle size and ζ-potential. Initially, the particle size distributions of CPI-stabilized emulsions were assessed by laser particle sizing at pH 7.0 (Fig. 5A), Native CPI emulsions exhibited a bimodal particle size distribution, with peaks exceeding 1000 nm, which were identified as oil droplets devoid of adsorbed proteins (Zhang, Zhao, et al., 2023). This non-uniform dispersion likely resulted from low electrostatic repulsion, which ultimately led to droplet aggregation and increased particle size. The particle size distribution remained bimodal after single modifications. Notably, the average particle sizes of U-CPIE and G-CPIE decreased from 7.04 in the unmodified CPI to 6.19 and 6.05 μm, respectively. This may be attributed to the cavitation effect generated by ultrasonic treatment during the protein modification stage, which altered protein conformation and weakened interparticle aggregation forces (Li, Xi, et al., 2022), resulting in reduced particle size and slightly altering the ζ-potential (Fig. 5B). In G-CPIE, the covalent connection of FG introduced spatial steric resistance and enhanced electrostatic repulsion, which was proved by the increase of the absolute value of ζ-potential (Li et al., 2021).

Fig. 5.

Particle size distribution (A), ζ-potential (B), adsorbed protein (C), and apparent viscosity (D) of different modified CPI stabilized emulsions.

Following dual modifications, the particle size showed a unimodal distribution, with U-G-CPIE reduced to 4.22 ± 0.02 μm, indicating that the ultrasonic treatment effectively dispersed the particles, by carrying out the glycosylation reaction, the proteins were able to stabilize the small oil droplets in the emulsion more effectively through hydrophilic-hydrophobic interactions (Wang et al., 2019). Additionally, the high negative charge of the U-G-CPIE (−47.93 mV) indicated the presence of strong electrostatic repulsion between the particles foaming the emulsion, leading to the effective prevention of coalescence. To summarize, the particle size distribution and ζ-potential of proteins in emulsions influence the stability of emulsions. These results align with recent studies that whey proteins achieved high ζ-potential with multi-scale ultrasound and polysaccharide addition.

3.4.2. Adsorbed protein

AP% represents the adsorption efficiency of proteins on the surface of emulsified oil droplets. It serves as a pivotal indicator of the ability of protein to stabilize the emulsion by covering the oil-water interface (Diao et al., 2024). As illustrated in Fig. 5C, the AP% of G-CPIE (71.66 %) was markedly higher than that of CPIE (62.01 %). This increase can be attributed to the interaction between plant proteins and anionic polysaccharides, with the polysaccharides promoting the continued adsorption of proteins on the surface of oil droplets by increasing repulsion between negative charges (Zhang et al., 2022). Notably, after dual modification, the AP% increased further to 84.56 % for U-G-CPIE and 77.36 % for G-U-CPIE. The higher AP% in U-G-CPIE suggests that sonication first altered the spatial arrangement of the protein structure and increased its specific surface area, while subsequent glycosylation enhanced the solubility and ζ-potential, allowing the smaller protein molecules to diffuse more easily to the oil droplet surface, further increasing the AP%.

3.4.3. Apparent viscosity

The relationships between apparent viscosity and shear rate for various emulsions are illustrated in Fig. 5D. All emulsions exhibited reduced viscosity as the shear rate increased. CPI is a globular protein (Fig. 3), and its viscosity decreases at low shear rates. This may be attributed to weak interactions between particles, which can be disrupted under low shear conditions, resulting in low viscosity (Yang et al., 2023; Yin et al., 2015). The slight increase in apparent viscosity observed following ultrasonic modification can be attributed to the reduction in the emulsion particle size, as smaller particles typically lead to higher viscosity. G-CPIE, U-G-CPIE, and G-U-CPIE displayed higher viscosities, with U-G-CPIE showing the highest. This increase can be attributed to the unfolding of the protein structure induced by ultrasound, creating additional binding sites. The subsequent addition of FG, which exhibits high thickening and gelling properties, markedly increased the apparent viscosity, consistent with the ζ-potential results.

3.5. Microstructural analysis of emulsions

The microstructures of emulsions stabilized by differently treated CPI fractions were examined using CLSM (Fig. 6). The control emulsion showed substantial aggregation and flocculation of oil droplets, likely due to insufficient protein adsorption at the oil-water interface. To alleviate this issue, different modification treatments were implemented. Both the single sonication and glycosylation treatments reduced flocculation, as previously observed by Xia et al. (2024) in PPI-stabilized emulsions, resulting in a finer particle distribution. In contrast, the dual-modified CPIEs demonstrated superior dispersion and homogeneity in droplet size; this was particularly evident in U-G-CPIE, which showed an increase in smaller particles and a finely distributed appearance. These findings were in agreement with the results of the droplet size and ζ-potential measurements, confirming the enhanced interfacial stabilization of U-G-CPIE.

Fig. 6.

CLSM images of different modified CPI-stabilized emulsions. CPIE (A), U-CPIE (B), G-CPIE (C), U-G-CPIE (D), and G-U-CPIE (E).

3.6. Stability of emulsions against environmental stresses

3.6.1. Thermal stability

Thermal stability is crucial for the application of substances in the food industry. In this study, we investigated the effect of heating on the particle size distribution of CPIEs. Untreated CPIE showed a significant increase in particle size after heating, primarily due to heat-induced disruption of the interfacial protein network, which weakened its ability to stabilize oil droplets. Additionally, as demonstrated in the studies by Wang et al. (2019) and Li, Zhang, et al. (2022), heating can promote protein unfolding and aggregation, increase surface hydrophobicity, and lead to droplet coalescence. In contrast, U-G-CPIE and G-U-CPIE showed minimal changes in particle size after heating. The combined effects of sonication and glycosylation enhanced both elasticity and stability of CPI during the heating phase. Additionally, the modified proteins were capable of rapidly restructuring the protective layer around oil droplets, preventing agglomeration and maintaining stable droplet sizes.

3.6.2. Ionic strength stability

To further investigate the suitability of the emulsions for practical applications at different salt concentrations, salt ion stability was assessed. As shown in Table 2, no significant change in the particle sizes in the emulsions was observed at a concentration of 150 mM, indicating that low salt concentrations had no adverse effects on emulsion stability. However, at 300 mM, particle size increased significantly and the stability of the emulsion system was impaired. The introduction of salt ions into the emulsion altered the surface charge of the emulsion, subsequently reducing the strength of the electrostatic forces between the emulsions, and thereby facilitating their aggregation. In contrast, after the dual modification, U-G-CPIE, especially, showed increased resistance to the presence of salt ions, with the particle size decreasing after the addition of salt ions, suggesting increased spatial repulsion, as has been observed in egg white protein (Wang et al., 2019), and pea protein isolates (Zhang, Wang, et al., 2023).

Table 2.

The effects of thermal, NaCl concentration, and storage time on the particle size distribution of emulsions with different modifications.

Condition		CPIE	U-CPIE	G-CPIE	U-G-CPIE	G-U-CPIE
Heating temperature (°C)	0	7.04 ± 0.77a	6.19 ± 0.06b	6.05 ± 0.23b	4.22 ± 0.02d	4.98 ± 0.01c
	90	12.70 ± 0.07a	7.19 ± 0.17a	7.18 ± 0.16b	4.48 ± 0.05c	5.25 ± 0.17bc
NaCl concentration (mM)	0	7.04 ± 0.77a	6.19 ± 0.06b	6.05 ± 0.23b	4.22 ± 0.02d	4.98 ± 0.01c
	100	8.12 ± 0.38a	6.42 ± 0.05b	7.18 ± 0.17c	4.21 ± 0.03e	4.75 ± 0.04d
	300	10.60 ± 0.70a	7.05 ± 0.25b	7.61 ± 0.18b	4.18 ± 0.01c	4.95 ± 0.34c
Storage duration (d)	0	7.04 ± 0.77a	6.19 ± 0.06b	6.05 ± 0.23b	4.22 ± 0.02d	4.98 ± 0.01c
	7	8.26 ± 0.63a	6.48 ± 0.09b	6.28 ± 0.19b	4.19 ± 0.07c	5.02 ± 0.00d
	14	9.06 ± 0.13a	7.01 ± 0.07b	6.94 ± 0.20b	4.25 ± 0.01c	5.05 ± 0.03d

Note: Canola protein isolate emulsion (CPIE), Ultrasound-canola protein isolate emulsion (U-CPIE), Glycation-canola protein isolate emulsion (G-CPIE), Ultrasound-glycation-canola protein isolate emulsion (U-G-CPIE), Glycation-ultrasound-canola protein isolate emulsion (G-U-CPIE), Different lowercase letters in the same column indicated significant differences (P < 0.05).

3.7. Storage stability of emulsions

Changes in the mean particle size were observed over 14 days (Table 2). The considerable enlargement of the CPIE particles following prolonged storage indicated poor stability, which was enhanced to varying extents by both sonication and glycosylation treatments. The particle size of U-G-CPIE and G-U-CPIE remained almost constant over storage, indicating enhanced stability during storage without significant external forces. This stability may be due to the following potential reasons: 1) ultrasound promotes the unfolding of the CPI structure and accelerates the glycosylation reactions through the interaction of aliphatic amino acids, thereby preventing emulsification or delamination of the emulsion (Zhang et al., 2024); 2) The synergistic effect of sonication and glycosylation leads to stronger repulsion between oil droplets on long-term storage, intensifying friction between the droplets and thereby preventing emulsion agglomeration and phase separation (Wang et al., 2022).

4. Conclusion

In this study, canola protein isolate was modified using a combination of glycosylation and ultrasound treatments. Ultrasound treatment effectively altered the protein structure, promoting the formation of additional disulfide bonds and enhancing hydrophobic interactions. The Maillard reaction following the addition of FG further enhanced the solubility, water-holding capacity, oil-holding capacity, emulsifying activity, and emulsion stability of the protein. Consequently, the double-modified U-G-CPI and G-U-CPI-stabilized flaxseed oil emulsions demonstrated superior physical properties. In particular, U-G-CPIE was associated with smaller droplet sizes, higher ζ-potential, and greater interfacial adsorption, resulting in the best stability during storage and application. These improvements effectively address the limitations of emulsions stabilized by CPI, U-CPI, and G-CPI. In conclusion, the combination of glycosylation and ultrasonication protein modifications represents a promising strategy, and these findings will contribute to the development of CPI emulsions and the effective expansion of CPI applications in the food industry.

CRediT authorship contribution statement

Ziqin Ye: Writing – review & editing, Writing – original draft, Methodology, Formal analysis. Jinying Wang: Writing – review & editing, Writing – original draft, Resources, Project administration, Methodology, Conceptualization. Guilan Ma: Methodology, Formal analysis. Jinge Ma: Methodology, Formal analysis.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Data availability

All data generated or analyzed during this study are included in this published article. Data will be made available on request.